POPULATION DYNAMICS OF RHIZOBIUM JAPONICUM AND RHIZOBIUM … II.pdf · POPULATION DYNAMICS OF...

POPULATION DYNAMICS OF RHIZOBIUM JAPONICUM

AND RHIZOBIUM LEGUMINOSARUM IN HOST

AND NON-HOST RHIZOSPHERES

A THESIS SUBMITTED TO THE GRADUATE DIVISION OF THE UNIVERSITY OF HAWAII IN PARTIAL FULFILLMENT

OF THE REQUIREMENTS FOR THE DEGREE OF

MASTER OF SCIENCE

IN AGRONOMY AND SOIL SCIENCE

DECEMBER 1982

By

Robert Baines Woolfenden II

Thesis Committee:

B. Ben Bohlool, Chairman Mitiku Habte Jake Halliday

We certify that we have read this thesis and that in our opinion

it is satisfactory in scope and quality as a thesis for the degree of

Master of Science in Agronomy and Soil Science.

THESIS COMMITTEE

.. ^ ^ Chairman

ACKNOWLEDGEMENTS

I am very grateful to Dr. B. Ben Bohlool for his guidance,

constructive criticisms, enthusiasm, and patience throughout all

phases of this research.

In addition, I am grateful to fellow graduate students,

Mark Kingsley, Renee Kosslak, Michael Sadowsky, and Paul Singleton,

for helpful suggestions and encouragement as the experiments pro

gressed.

This work was supported in part by Grant AID/ta-C-1207 (NifTAL

Project) from the United States Agency for International Development.

The conclusions reached within this study do not necessarily represent

the views of the granting agency.

TABLE OF CONTENTS

Page

ACKNOWLEDGEMENTS......................................... iii

LIST OF TABLES ......................................... v

LIST OF F I G U R E S ......................................... vi

LIST OF DIAGRAMS......................................... viii

CHAPTER 1. INTRODUCTION AND LITERATURE SURVEY ............ 1

CHAPTER 2. POPULATIONS OF RHIZOBIUM IN THE RHIZOSPHERESOF HOST AND NON-HOST PLANTS AT HARVEST......... 17

CHAPTER 3. POPULATIONS OF RHIZOBIUM IN THE RHIZOSPHERES OF HOST AND NON-HOST PLANTS DURING A 35-DAY GROWTH CYCLE....... ........................... 47

CHAPTER 4. RHIZOBIUM POPULATION DYNAMICS IN HOST AND NON-HOST RHIZOSPHERES DURING THE FIRSTNINE DAYS OF RHIZOSPHERE DEVELOPMENT........... 85

iv

CHAPTER 5. GENERAL DISCUSSION 107

V

Page

TABLE

2-1 Comparison of the recovery of added Rhizobiumfrom Kula loam soil by two methods.................. 30

4-1 Percentages of Rhizobium inoculant strains(combined) in the total rhizosphere populationof bacteria at 3, 5, 7, and 9 days.................. 100

4-2 Percentages of Rhizobium inoculant strains(combined) in the total rhizosphere populationof bacteria at planting, 24, and 48 hours........... 101

LIST OF TABLES

vi

££

31

32

33

34

35

36

38

39

39

59

60

61

62

63

64

65

66

LIST OF FIGURES

Counts of 4 Rhizobium strains at harvest 1




Acridine orange total counts of bacteria at harvests 2, 3, and 4..................

Soybean nodule occupancy by Rhizobium japonicum strains........................

Pea nodule occupancy by Rhizobium leguminosarum strains....................

R. japonicum nodule bacteria, stained with homologous fluorescent antibody . . .

R. 1eguminosarum nodule bacteria, stained with homologous fluorescent antibody . . .

Counts of 4 Rhizobium strains per gram of soil at 1 week ..........................

Counts of 4 Rhizobium strains per gram of soil at 2 weeks.........................


Counts of 4 Rhizobium strains per gram of soil at 4 weeks..........................


Counts of 4 Rhizobium strains per square centimeter of root area at 1 week........

Counts of 4 Rhizobium strains per square centimeter of root area at 2 weeks . . . .

Counts of 4 Rhizobium strains per square centimeter of root area at 3 weeks . . .

Page

FIGURE

3-9 Counts of 4 Rhizobium strains per squarecentimeter of root area at 4 weeks................ 67

3-10 Counts of 4 Rhizobium strains per squarecentimeter of root area at 5 weeks................ 68

3-11 Acridine orange total counts of bacteria pergram of soil over 5 weeks........................ 69

3-12 Acridine orange total counts of bacteria persquare centimeter of root over 5 weeks. ........ 70

3-13 Soybean nodule occupancy by R. japonicumstrains......................................... 72

3-14 Large pea nodule occupancy by R. leguminosarumstrains......................................... 73

3-15 Small pea nodule occupancy by R. leguminosarumstrains....................... 74

3-16 View of the experiment at 1 week................ . 75

3-17 View of the experiment at 3 weeks................ 77

3-18 View of the experiment at 5 weeks................ 77

4-1 Counts of 4 Rhizobium strains at 3 days........... 92



4-4 Counts of 4 Rhizobium strains at 9 days ........... 95

4-5 Acridine orange total counts of bacteriaat 3, 5, 7, and 9 days............................ 96

4-6 Counts of 4 Rhizobium strains in the pearhizosphere at planting, 24, and 48 hours......... 97

4-7 Counts of 4 Rhizobium strains in the soybeanrhizosphere at planting, 24, and 48 hours......... 98

vii

LIST OF FIGURES (Continued)

viii

FIGURE

4-8

LIST OF FIGURES (Continued)

Page

Acridine orange total counts of bacteria in the pea and soybean rhizospheres at planting,24, and 48 hours................................... 99

ix

Page

DIAGRAM

3-1 Positioning of watering device, and setup ofexperimental pot for watering from below.......... 54

3-2 Arrangement of cores and order of harvest forthe 35-day time course study...................... 55

LIST OF DIAGRAMS

CHAPTER 1

INTRODUCTION AND LITERATURE SURVEY

INTRODUCTION AND LITERATURE SURVEY

The increasing disparity between population growth and food

production has placed a burden on agriculturalists to devise more

efficient methods of food production. Therefore, the Rhizobium-

legume symbiosis has received renewed attention lately, as it becomes

more and more apparent that we cannot count on cheap fossil fuels to

supply our needs for fixed nitrogen. There has been a renascent

interest in biological nitrogen fixation in general, and in the

Rhizobium-legume symbiosis in particular, as a means to help provide

protein to the world's rapidly-expanding population.

The symbiosis is a partnership between soil bacteria of the

genus Rhizobium and plants of the family Leguminosae. The visual

manifestations of the symbiosis are legume root nodules which house

the rhizobia. Inside these nodules atmospheric dinitrogen gas is

enzymatically reduced initially to ammonia, thereby entering the

assimilation pathways of plants.

The term "rhizosphere" was coined by Hiltner in 1904 as "the

region of contact between soil and root, where the soil is affected

by the root".

Before any nodules are formed, however, a series of intricate

events occurs in the rhizosphere. Rhizobia multiply in the host

rhizosphere, somehow recognize the plants' roots as those of a host

legume, infect those roots, and eventually initiate nodule formation.

Specificity is the hallmark of the Rhizobium-legume symbiosis; only

certain species of rhizobia are able to infect the roots of partic

ular legume species. Although the relatioship between the bacteria

and the host legume has been studied in great detail, as yet the

basis for this specificity is not clear.

A number of authors (Bohlool and Schmidt, 1974; Dazzo and

Hubbel, 1975) have implicated lectins (sugar-specific proteins or

glycoproteins on the legume root surface) as a possible basis for

specificity. However, due to the lack of standardized techniques

for examining lectin binding, the notion of lectin-mediated speci

ficity is still controversial (Broughton, 1978, Schmidt, 1979).

Another possible contributor to specificity is the stimulation

of the growth of rhizobia in the rhizosphere of their homologous

host plants, over and above that of the normal rhizosphere flora,

and over other non-homologous rhizobia (Nutman, 1963, 1965). This

specific stimulation in the rhizosphere has been postulated to be

mediated by root exudates. Selective stimulation of rhizobia by the

roots of their legume hosts would be of great selective advantage to

both, and presumably could be operating in concert with lectins to

confer specificity to the symbiosis.

The objective of this chapter is to give a concise review of

the pertinent literature on the effects of root exudates on rhizobia,

and to examine specific stimulation by those exudates as a possible

contributor to specificity in the symbiosis.

Plant roots in general stimulate gram negative rods more than

other soil bacteria, and these constitute greater proportions of the

rhizosphere than the soil populations in general (Rovira and

McDougall, 1967). Krasil'nikov (1958) demonstrated that bacteria

which colonize the rhizosphere of one plant species do not necessarily

colonize that of other plant species. He also showed that not all

strains of Pseudomonas fluorescens have equal rhizosphere colonizing

abilities. One of the most consistent differences between bacteria

isolated from the rhizosphere and those isolated from fallow soil is

the requirement of the former for amino acids (Lochhead and Rouatt,

1955).

Starkey (1929) suggested the following factors to be involved in

the stimulation of microorganisms by plant roots; sloughed-off root

cells, moribund root hair and cortical cells, and soluble organic

nutrients released or actively exuded by intact roots. Much evidence

has been presented demonstrating the exudation of many organic materials

from healthy, intact plant roots (Scroth and Hildebrand, 1964; Rovira,

1962; 1965). These studies support the notion that sufficient

nutrients are exuded to support large rhizosphere populations of

microorganisms.

The legume rhizosphere has shown to be a particularly intense

zone of microbial activity, due to the nature and quantity of products

exuded by legume roots (Rovira, 1956a; 1962). According to the theory

of specific stimulation, legume exudates should stimulate rhizobia

able to infect them more than other rhizobia. This theory was detailed

by Nutman (1965).

Most of the early studies were conducted in aseptic solution,

agar, or sand culture, using one or a few strains of Rhizobium. The

following is a chronological review of some of the early studies.

Nutman (1953) grew clover, alfalfa, and vetch, singly and in

pairs consisting of either one or two plant species. Plants were

4

cultured aseptically in tubes on agar "slopes". In these experiments

the presence of an alfalfa plant in the same tube with a clover plant

and R. trifolii, plants were nodulated sooner than in tubes in which

there were two clover plants. Apparently, exudates from the alfalfa

roots were able to stimulate the nodulation of clover. Exudates

from inoculated clover plants uniformly inhibited the nodulation of

another clover, lucerne, or vetch plant in the same tube. This

inhibition was least in clover and greatest in vetch. Inhibition of

nodule formation on clover was obtained in tubes which had been

preplanted with clover and the earlier plantings removed. Nutman

interprets his results in terms of the secretion of some nodule-

inhibiting substance from the clover roots.

Several of the early examinations of plant root exudates in the

rhizosphere were carried out by Rovira (1956a, 1956b, 1961; Rovira

and Harris, 1961). In many of his studies, plants were cultured in

sterile sand, -instead of the sterile water or agar culture used by

several workers previously. Following growth, plants were carefully

removed, and exudates leached out of the sand. Rovira (1956a)

demonstrated the exudation of a host of amino compounds and sugars

from aseptically-grown oat and pea plants. Exudates from pea roots

were larger in quantity, and compounds were more numerous and varied

than those from the roots of oats. Early studies had been complicated

by the fact that only a small amount of rhizosphere soil was available

for study. To get around this problem, Rovira (1956b) established an

"artificial rhizosphere" in which non-sterile field soils were

saturated with root exudate solution collected from aseptically-grown

5

roots. Treatment was continued for 21 days, and an overall

stimulation of bacterial growth upon the addition of the exudates,

the bacteria consisting largely of gram negative rods, was taken as

evidence that his artificial rhizosphere had actually been successful.

The resulting populations of root-exudate-treated soils did not

increase the rate of organic matter decomposition in the soil.

However, the artificial rhizosphere population did promote the

decomposition of the more readily-available organic nutrients, such

as amino acids and glucose. Rovira (1961) compared the rhizospheres

of red clover and paspalum (a grass) with respect to numbers of R.

trifolii and total bacteria. In this study, non-sterile field soil

was used, and Rhizobium numbers were determined by the most probable

number (MPN) technique (Vincent and Waters, 1954). Bacterial numbers

were examined in response to the rhizospheres of the two plant

species and also to varying lime levels. Bacteria, including

rhizobia, were consistently present in larger numbers in the rhizo

sphere of clover than that of paspalum; ratios for Rhizobium were

about 5:1 (clover: paspalum) and for total bacteria, about 2:1.

Liming was beneficial to the rhizosphere populations of both plants.

The liming probably affected the growth of the microorganisms

directly and also indirectly, by first enhancing plant growth and

root exudation. In a further study, Rovira and Harris (1961)

examined the exudation of growth factors from peas, alfalfa, tomato,

and several clover species grown in sterile sand culture. Their aim

was to quantitatively assess the various B-group vitamins exuded by

these plant species. Biotin was found to be exuded in the largest

6

quantity, and was found in the rhizosphere of pea at 10 to 100 times

the concentration found in clover or tomato rhizospheres. Other

growth factors, notably pantothenate and niacin, were present, but in

amounts considered by the authors as unlikely to influence the growth

of microorganisms. In non-sterile sand culture, biotin and panto

thenate were seen to disappear rapidly, emphasizing the need for

strict asepsis in any studies of root exudates.

Studies by Rovira and coworkers emphasized the large variety and

quantity of substrates exuded by plant roots, particularly those of

legumes. However, most of these studies focused on the exudates

themselves rather than the particular bacteria stimulated by those

exudates. Because of this fact, most studies were carried out under

aseptic conditions in sand culture.

Elkan (1961) examined a non-nodulating, near isogenic soybean

variety to determine the reason for non-nodulation. In greenhouse

solution culture, he demonstrated that the root excretions from the

mutant non-nodulating line resulted in highly significant (p=.01)

decreases in nodulation of normal, nodulating plants. In addition,

the excretion resulted in decreased total nodule weight, total dry

weight, and total nitrogen per nodulating plant. Curiously, the

excretion did not inhibit growth of R. japonicum directly, nor did

it inhibit nodulation of other plant species by other rhizobia.

Tuzimura and Watanabe (1962a) examined the populations of

bacteria, fungi, actinomycetes, and specifically Agrobacterium

radiobacter and a Rhizobium spp. from Astragalus sinicus in the

rhizosphere of this plant. A non-sterile volcanic ash soil was used,

7

and rhizobia were enumerated by an MPN method. The astragalus plants

were able to support a reasonable population of astragalus rhizobia,

about 8.1 x 10** at flowering, and 10® per gram dry root at fruiting,

regardless of starting Rhizobium population. In a further study,

Tuzimura and Watanabe (1962b) followed populations of R. trifolii in

the rhizospheres of several leguminous plants, as well as the rhizo

spheres of non-legumes. Ladino clover, alfalfa, soybean, and peanut

were the legumes examined. The non-legumes were sudan grass and

upland rice. Rhizobium trifolii were consistently seen in larger

numbers in the rhizosphere of alfalfa, followed by peanut, followed

by soybeans, and, followed by clover. No statistical treatment of

the data was presented, and the authors state that "these findings

might not be necessarily valid, because the growth of the plants and

contents of soils which adhered to the root varied in each case."

Rhizosphere numbers of R. trifolii were, without exception, at least

two logs greater in the rhizosphere of legumes than in non-legume

rhizospheres.

Nutman (1963, 1965) suggested that "A given legume tends to

promote the multiplication of bacteria able to infect it more than

others", and, "Individual strains of nodule bacteria are more

strongly stimulated by those hosts they are able to infect than by

other legumes", citing only a reference by Wilson (1930) in support

of these statements. Dart and Mercer (1964) present the opposite

viewpoint. They state that there is no evidence that legume roots

selectively stimulate the growth of Rhizobium rather than other

organisms. Further, they state that the Rhizobium strains which

8

nodulate a particular legume are not preferentially stimulated in that

hosts rhizosphere over other Rhizobium strains, citing Krasil'nikov

(1958) and Purchase (private communication).

Rovira (1965) concluded that sufficient chromatographic

analyses had been performed on root exudates to indicate the wide

spectrum of compounds contained therein.

Peters and Alexander (1966) examined four different rhizobia in

the rhizosphere of alfalfa and Lotus corniculatus in aseptic solution

culture. Alfalfa was inoculated with R.. meliloti, R. trifolii, and

R. leguminosarum to see if alfalfa stimulated only its homologous

Rhizobium, R. meliloti. Despite differences in initial inoculum

size and in spite of the fact that R. meliloti alone induced

nodulation, populations of all rhizobia reached roughly 106 to

107 cells/ml of rooting medium after 1 week, and following that cell

numbers did not fall rapidly. These results, albeit in aseptic

solution culture, were not suggestive of specific stimulation. Also,

when alfalfa and Lotus corniculatus were inoculated with a mixed

culture of their respective microsymbionts, no selective interaction

between host and homologous organisms was seen. Peters and Alexander

(1965) suggested that the selectivity between microbe and host is

probably exerted first at individual receptor sites on host root

surfaces.

Van Egeraat (1975a) examined the growth of R. leguminosarum on

the rhizoplane (root surface) and in the rhizosphere of aseptically

growth pea seedlings, using sterile agar culture in petri plates.

He observed no bacterial growth on the main taproot, however, after

9

the emergence of secondary roots bacterial numbers increased where

the secondary roots emerged from the main taproot. Lateral roots of

the pea seedlings growing in sterile agar along the bottom of the

petri dish had a zone of bacterial growth some distance from the

roots. Van Egeraat (1975a) attributed this to a zone of growth

inhibition surrounding roots more closely. Of particular interest

in this study was the stimulation of rhizobia around the sites of

emergence ("wounds") of the lateral roots. Van Egeraat concludes

that young pea plants exude both growth stimulating and growth-

inhibiting compounds. Further, the growth-inhibiting compounds can

temporarily prevent the growth of R. leguminosarum in the immediate

vicinity of the pea roots. In a follow-up study, Van Egeraat (1975b)

demonstrated that R. 1eguminosarum grew equally well with homoserine

and glutamic acid as the nitrogen source, or as the sole source of

carbon, nitrogen, and energy. Strains of R. trifolii, R. phaseoli,

and R. meliloti behaved entirely differently. These three strains

could grow with glutamate as the only C and N source. With homo

serine, growth was extremely slow or absent, or in the case of R.

meliloti, considerably reduced. Van Egeraat suggested that homo

serine (which was found to comprise about 70% of the amino compounds

exuded by pea roots) might selectively stimulate the growth of R.

leguminosarum in the pea rhizosphere when a mixture of Rhizobium of

many species is present. This represents perhaps the most compelling

study in favor of specific stimulation. However, it was carried out

under totally aseptic conditions, and on the basis of these experi

ments it would be impossible to predict the fate of homoserine in the

10

rhizosphere of pea plants grown in the field. Homoserine is the first

compound to be suggested as a specific stimulant in the scientific

literature. Rovira (1965) stated that it would appear unlikely that

the the ubiquitous sugars and amino acids would provide the speci

ficity observed in the symbiosis, but rather the balance of these

compounds of the presence of exotic compounds peculiar to a particular

plant species. Van Egeraat (1975b) states that in the pea system,

just such a compound, homoserine, is present.

The studies on root exudates and the rhizosphere effect with

respect to rhizobia have either addressed the interaction in aseptic

systems, from which organisms can be conveniently plated and counted,

or have enumerated rhizobia indirectly by means of MPN counts (in the

case of non-sterile systems). However, either method has its share

of disadvantages, and thus early studies of individual species or

strains of Rhizobium in the rhizosphere of their homologous host

plants are not without criticism. What was needed was an adequate

methodology for strain-specific enumeration of rhizobia in natural

non-sterile rhizospheres, in which the full range of microbe-to-microbe

and plant-to-microbe interactions are taking place.

An autoecological approach, in which the numbers of several

different Rhizobium strains in the rhizospheres of several host

plants could be studied, is necessary to adequately address the

problem. The only method adequate for the study of a specific micro

organism directly in a natural soil environment is immunofluorescence

(Schmidt, 1979; Bohlool and Schmidt, 1980).

11

Reyes and Schmidt (1979) used membrane filter immunofluorescence

(Schmidt, 1974) to enumerate R. japonicum strain 123 in soil and in

rhizospheres of Minnesota field-grown soybeans. Rhizosphere effects

were modest. About 101* to 105 cells/gram of soil were observed in

soil adhering to plant roots. A comparably slight rhizosphere effect

was seen for corn. According to their data, strain 123 did not

follow what they termed the "scenario" of specific stimulation.

Further, these authors state that experimental support for the

specific stimulation hypothesis is meager, especially in terms of

rhizobial response to plant rhizospheres under natural soil conditions.

In a further study, Reyes and Schmidt (1981) used membrane filter

immunofluorescence for enumeration of, and immunofluorescence

examination of strain USDA 123 on root surfaces of field-grown

soybeans. Strain 123 was a consistent rhizosphere colonizer, but

failed to multiply as rapidly as the root system developed under

field conditions. Root surface populations declined from about

7 to 8 x 102 per square cm of root surface on day 9 to about 60 per

square cm on day 26. Average calculated cell density per square cm

was only a few hundred. They also examined the behavior of strain 123

in the rhizospheres of pot-grown soybeans. In this trial, cell

numbers and root surface populations were monitored every 4 days,

until nodule initiation. This greenhouse experiment confirmed the

findings of the field experiment. Strain 123 was seen in numbers of

a few hundred per square cm of root surface during early growth, and

declined to less than 100 per square cm with more extensive root

development. Also addition of a competing strain (USDA 138) had

12

little effect on the development of strain 123 in the soybean

rhizosphere. Even when 10 times as many 138 cells than 123 cells

were added, cell densities of both stabilized at a few hundred per

square cm of root surface, and each appeared to establish indepen

dently of the other. Immunofluorescence examination of unwashed

roots revealed that both strains were sparsely distributed on root

surfaces. Microscopic examination of the roots also revealed that

rhizobia were seldom seen in microcolonies, but rather as single,

double, or triple cells. In this study, no proliferation of

Rhizobium was observed, even at the junction of lateral and tap roots

where release of organic compounds is likely to occur due to the

wounds caused by the emergence of secondary roots (Van Egeraat,

1975a). No evidence was obtained in support of specific stimulation

as a prelude to nodulation. Strain 123 established in numbers

roughly equal to those of 138, even though 123 forms the vast

majority of the nodules. Performing rhizosphere counts on the basis

of root surface area makes the lack of specific stimulation in the

rhizosphere even more clear cut. Competitive advantages of

strain 123 over strain 138 were not obvious, however. These studies

by Reyes and Schmidt represent an important first step in examining

rhizobia in natural non-sterile field soil rhizospheres.

Exudates are important in establishing and maintaining the

rhizosphere population of microorganisms; bacteria in particular

seem to respond to the soil conditions around plant roots. In

aseptic systems, the interactions between exudates and bacteria

(particularly Rhizobium) have been studied by many investigators.

13

Results in sterile systems have been somewhat ambiguous. In

autoecological studies done under natural field soil conditions, no

evidence has to date been found in support of specific stimulation.

However, no study has examined several different species of rhizobia

in the rhizospheres of both homologous and non-homologous legumes.

An appropriately-designed autoecological study such as this could

adequately address the question of specific stimulation in the legume

rhizosphere.

The basis for specificity in the Rhizobium-legume symbiosis is

still a matter of debate. A better knowledge of this basis might

ultimately help to extend nodulation and nitrogen fixation capability

beyond the legumes to other grain crops, grasses, and cereals. Self-

sufficiency for nitrogen would be a highly desirable trait for any

crop plant.

14

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24. Rovira, A. D. and B. M. McDougall. 1967. Microbiological and biochemical aspects of the rhizosphere. pp. 417-463. InA. D. McLaren and G. F. Peterson (Eds.) Soil biochemistry. Marcel Dekker, New York.

25. Schmidt, E. L. 1974. Quantitative autoecological study of microorganisms in soil by immunofluorescence. Soil Sci. 118: 141-149.

17

26. Schmidt, E. L. 1979. Initiation of plant root-microbeinteractions. Ann. Rev. Microbiol. 33:355-376.

27. Scroth, M. N. and D. C. Hildebrand. 1964. Influence of plant exudates on root-infecting fungi. Ann. Rev. Phytopathology 2: 101-132.

28. Starkey, R. L. 1958. Interrelations between microorganisms andplant roots in the rhizosphere. Bact. Rev. 22:154-168.

29. Tuzimura, K. and I. Watanabe. 1962a. The growth of Rhizobium in the rhizosphere of the host plant. Ecological studies of root nodule bacteria (part 2). Soil Sci. Plant Nutri. (Tokyo) 8:19-24.

30. Tuzimura, K. and I. Watanabe. 1962b. The effect of various plants on the growth of Rhizobium. Ecological studies of root nodule bacteria (part 3). Soil Sci. Plant Nutri. (Tokyo) 8:13-17.

31. Vincent, J. M. and L. M. Waters. 1954. The root nodule bacteriaas factors in clover establishment in the red basaltic soils of the Lismore district, New South Wales. II. Survival and success of inocula in laboratory trials. Aust. J. Agric. Res. 5:61-76.

32. Wilson, J. K. 1930. Seasonal variation in the numbers of two species of Rhizobium in soil. Soil Sci. 30:289-296.

POPULATIONS OF RHIZOBIUM IN THE RHIZOSPHERES

OF HOST AND NON-HOST PLANTS AT HARVEST

CHAPTER 2

ABSTRACT

The growth of two strains of R. japonicum (strains USDA 110 and

CB 1809), and two strains of R. leguminosarum (strains Hawaii 5-0 and

Nitragin 92A3) was followed in the rhizospheres of soybean, pea, and

corn growing in non-sterile soil. Rhizosphere soil was sampled at

35 days over four successive growth cycles. The numbers of each strain

were determined by membrane filter immunofluorescence, using strain-

specific fluorescent antibodies. Nodule occupancy of the strains on

their appropriate host was determined by immunofluorescence. No

specific stimulation of rhizobia in the rhizospheres of their homologous

host plants was observed. Counts of all strains, as well as total

bacteria were generally in the following order: soybean rhizosphere>

pea>corn>fallow soil. Strain CB 1809 occupied a slightly higher

percentage of soybean nodules than USDA 110, whereas Nitragin 92A3

dominated Hawaii 5-0 in pea nodules. Fifteen percent of soybean

nodules were doubly infected, as were 3.5% of pea nodules. The four

strains together comprised 1.5 to 2.6% of the total rhizosphere

bacteria of the legumes and 2 to 9% of the total bacteria in the corn

rhizosphere. Rhizobia comprised 3 to 5% of the total bacteria in

fallow soil. These data are not suggestive of an overwhelming increase

of homologous rhizobia in the rhizospheres of their respective host

legumes. Specific stimulation of growth of Rhizobium in the legume

rhizosphere does not appear to be a contributor to specificity in the

Rhizobium-legume symbiosis under the conditions of this study.

INTRODUCTION

Soil bacteria of the genus Rhizobium are recognized by their

ability to form nitrogen-fixing nodules on the roots of many leguminous

plants. Rhizobia are specific with respect to the hosts they nodulate.

Only certain species of rhizobia are able to infect and nodulate the

roots of particular legumes. Although the relationship between rhizobia

and their host legumes has been studied in great detail, the basis for

this specificity is not completely clear (5,20).

Several possible mechanisms have been advanced to account for

specificity, involving both plant and bacterial components (5).

Bacterial chemotaxis might affect rhizosphere populations, as might

rhizosphere competence once rhizobia have arrived in the root zone

(21). Cellular recognition between host and bacterium is considered

by some as a mechanism for specificity. Plant proteins called lectins

are thought to interact specifically with rhizobia at the root surface

(3,9,22). Another proposed mechanism for specificity is the prefer

ential growth of the appropriate rhizobia in the rhizospheres of their

homologous host plants over and above that of other rhizobia (7,14,25).

Most studies of stimulation in the legume rhizosphere have been

carried out in aseptic systems, from which rhizobia can be conveniently

plated and counted (8,13,15). However extrapolation from aseptic

conditions to non-sterile field soil, in which the full range of

microbial interactions would be occurring, is unrealistic, and often

erroneous.

An indirect method of enumerating rhizobia in non-sterile soil is

the plant dilution assay (24). In this method, the plant itself is

used as a selective agent, and rhizobia enumerated by dilution-

extinction. This method is cumbersome, however, and does not lend

itself to ecological studies.

Direct study of specific bacterial strains in soil and rhizo

spheres has been prevented by the lack of an adequate methodology for

enumeration of bacteria in the particulate soil environment (4).

Bohlool and Schmidt (2) and Schmidt (19) solved many of the method

ological problems with the introduction of a quantitative membrane

filter immunofluorescence (MFIF) technique for enumeration of specific

microorganisms directly in soil. This work was extended by Kingsley

and Bohlool (11) who adapted the MFIF technique for use in tropical

soils. In the present report, MFIF is used to examine the population

dynamics of homologous and non-homologous rhizobia in the rhizospheres

of host and non-host plants.

21

MATERIALS AND METHODS

Preliminary Soil Examination

Kula loam soil, an Inceptisol (Typic Eutrandept, pH 6.5), was

chosen as the experimental soil. The soil was tested for the presence

of cross-reactive bacterial cells, fungal spores, and mycelia by the

membrane filter immunofluorescence (MFIE) method of Kingsley and

Bohlool (11). Kula soil was also tested for the presence of

indigenous R. japonicum and R. leguminosarum by inoculating soybean

and pea seedlings grown in flasks of sterile vermiculite. Seeds were

surface sterilized by shaking in a 4% solution of sodium hypochlorite

for 20 minutes followed by at least five rinses in sterile water, and

were germinated on .9% water agar. One gram of Kula soil was added

directly to the radicles of aseptically grown 2- to 3-day-old soybean

or pea seedlings. Plants were grown in a growth chamber for 28 days,

and roots checked for nodulation.

Rhizobium Strains

R. japonicum strains CB 1809 and USDA 110 and R. leguminosarum

strains Hawaii 5-0 and Nitragin 92A3 were obtained from the collection

of B. B. Bohlool of the University of Hawaii. Strains were maintained

on yeast-extract mannitol agar slants (24), with 1.0 g of yeast extract

(Difco Laboratories, Detroit, Michigan) substituted for yeast-water.

Experiments to Assess Recovery

Kula soil was inoculated with known numbers of rhizobia, and their

recovery assessed by the method of Kingsley and Bohlool (11), using

appropriate FAs.

Inoculum Preparation

Yeast-extract mannitol broth (24) cultures of each of the four

strains were grown to early stationary phase. Kula loam soil (250 g)

was amended to 1% with mannitol, adjusted to about -1/3 bar moisture

tension with distilled water, and autoclaved (121 C, 15 psi) for

45 minutes on two successive days. Ten ml aliquots of the above broth

cultures were then added aseptically, and these soil cultures incubated

at room temperature for 3 days (R. leguminosarum) or 8 days (R.

japonicum). MFIF counts were performed on these soil cultures and

aliquots of each added to 1 kg of non-sterile Kula soil to a level of

about 5 x 105 cells of each strain, per gram of moist soil. This

mixture was homogenized by manual shaking in a large plastic bag for

10 minutes. MFIF counts were performed on this intermediate dilution

of inoculant. The intermediate dilution was mixed with an additional

4 kg of non-sterile soil to give about 1 x 10s cells per gram (moist

soil) of each strain. Final mixing was done in a twin-shell dry

blender (The Patterson-Kelley Co., Inc., East Stroudsburg, Pa.) for

10 minutes.

Pots and Planting

Pots used were 1400 ml polypropylene enema containers (Resiflex,

hospital surplus) painted with a heavy coat of flat white paint. Each

pot contained 750 grams of non-sterile inoculated Kula soil (moist

weight). Soils were brought to about -1/3 bars of tension with

distilled water. Three-day-old seedlings of peas (wilt-resistant

Wisconsin Perfection), soybeans (Davis), and corn (Hawaiian Supersweet

23

#9) were planted. Seeds were pregerminated aseptically on .9% water

agar for 3 days. Three corn plants, four soybean plants, and five pea

plants were sown into their respective pots. Pots were planted in

triplicate. Three pots were set up as fallow controls (non-rhizosphere

soil). Following planting, the soil surface was covered with rinsed

white aquarium gravel (California Wonder Rock, Kordon Co., Hayward,

Ca.) to prevent undue soil heating and to inhibit the growth of algae

on the soil surface. The experiment was set up as a randomized

complete block.

Plant Growth Conditions

Harvest cycles were of approximately 35 days. The experiment

was not run longer because the release of bacteroids at nodule

senescence would certainly give the appearance of rhizosphere stimulation

of specific rhizobia (23) . Soil moisture tension was maintained at

about -1/3 bars by daily watering to a constant weight with distilled

water. Hoagland's N-free medium (10) was substituted for water once a

week. For the first harvest, plants were grown in a climate controlled

plexiglas house, onto which was attached a large air conditioner to

maintain temperatures of 27°C during the day and 21 C at night.

Subsequent cropping cycles were carried out in a greenhouse, where

temperatures were slightly higher. Four cropping cycles were performed.

Following a harvest, soil from the preceding harvest was placed back

into the same pots without further amendment, and the pots replanted

with the same crop.

24

Harvest and Sample Preparation

At harvest, the entire contents of each pot was carefully removed

and the loosely adhering soil gently shaken free of the root system.

Any large adhering soil clumps were also removed. Root systems with

adhering soil were then placed in 18-ounce Whirlpak bags for transport

to the laboratory. The root systems with rhizosphere soil (including

that which rubbed off the roots into the bags) were transferred to

square, wide mouth screwcap bottles (45x45x120 mm, approximate

capacity 230 ml). To these root and rhizosphere soil samples was

added 100 ml of .1% partially hydrolyzed gelatin (diluted in .1 M

dibasic ammonium phosphate, 11). Four drops of tween 80 (Sigma) were

added, and the bottles tightly capped and shaken on a Burrel Wrist-

Action shaker for 30 minutes at full power. Immediately after shaking,

25 ml of suspension was removed from each bottle and transferred to a

50 ml polycarbonate centrifuge tube. Samples were centrifuged gently

(700 x g, 5 min.) in a Sorvall centrifuge, to pellet soil particles.

Following centrifugation, supernatants were decanted into clean

screwcap tubes. The soil pellet, as well as the remaining soil and

soil solution was then rinsed into a pre-weighed aluminum tart pan for

soil dry weight determination. Pans were held at 105°C for 48 hours,

and weighed immediately after having cooled to room temperature. The

weight of the dried gelatin-ammonium phosphate preparation was deter

mined experimentally, and this value subtracted from soil weights.

Non-rhizosphere soil was examined using a modification of the

Kingsley/Bohlool MFIF procedure (11). Ten grams of non-rhizosphere

soil was placed in a 250 ml screwcap Erlenmeyer flask, and 1/3 of a

25

standard scintillation vial full of 3 mm diameter glass beads added.

To this was added 10 ml of 10% hydrogen peroxide (E. L. Schmidt,

personal communication). This mixture was shaken on a Burrel Wrist-

Action shaker for 10 minutes at full power. Flasks were capped some

what loosely, to prevent explosion. Following this step, 75 ml of

gelatin-ammonium phosphate (11) was added, and the mixture shaken for

an additional 20 minutes. Twenty-five ml of soil suspension was

removed from the flasks, transferred to 50 ml polycarbonate centrifuge

tubes, and centrifuged as above. Rhizosphere and non-rhizosphere

samples were diluted 1:10 and 1:2, respectively, before filtration for

FA counting.

Membrane Filter Immunofluorescence

Membrane filter counts were performed as in Kingsley and

Bohlool (11). Following filtration of the sample, filters were placed

on microscope slides, and the effective filtering area covered with

six drops of partially hydrolyzed gelatin-rhodamine isothiocyanate

conjugate (1). This served to reduce non-specific adsorption of

fluorescent antibody to soil colloids, and control background

fluorescence. RhITC-gelatin treated filters were then dried at 50°C

and held at that temperature until staining with the fluorescent

antibody. Filters were stained for 1 hour. Following staining,

filters were placed back on filter holders and immediately rinsed

with at least 150 ml of prefiltered .85% saline, then returned to

microscope slides, mounted with a coverslip in buffered glycerol

(pH 9), and observed. Microscopy was performed with a Zeiss standard

microscope 14 equipped with incident light illumination from an HBO 50

26

(Osram) mercury vapor light source and a Zeiss fluorescein

isothiocyanate (FITC) filter pack. A Zeiss 63X Planapo oil immersion

objective was used. Duplicate filters were counted for each strain

from each pot, for a total of 96 filters per harvest. No correction

factor was used to account for the fact that recoveries are generally

about 70 to 95%.

Acridine Orange Total Counts

Estimations of the total rhizosphere population of bacteria were

made at each harvest using acridine orange. Dilutions of centrifuge

supernatants were made in water which was collected directly from a

glass still into an acid-washed flask. Samples were concentrated onto

membrane filters, and filters transferred to microscope slides.

Filters were stained directly on the slides with one drop of acridine

orange solution (Sigma, 1:30 000 in .1 M phosphate buffered saline

pH 7.2) previously filtered through a .2 micron membrane filter

(Gelman Acrodisc, Gelman, Ann Arbor, Mich.). Three filters were

counted per pot, for a total of 36 per harvest.

Nodule Typing

Following the extraction of the rhizosphere soil from roots, all

nodules were removed, squashed with forceps, and smeared on slides

for serological typing with fluorescent antibodies (18). Microscopy

was performed with incident light fluorescence illumination in

conjunction with transmitted-light phase contrast lighting.

27

Immunofluorescent Examination of Root Surfaces

Root surfaces were examined by immunofluorescence (6) to insure

that rhizobia bound to root surfaces were being released into suspension

for counting.

Photographs (Figs. 7 and 8) were made with Ektachrome 200 slide

film, and prints made from these slides.

28

RESULTS

The Kula soil was found to be a good experimental soil for the

application of immunofluorescence to the study of ecology of Rhizobium.

Recoveries of added inoculant rhizobia were consistently in the range

of 70 to 95% (Table 1). The hydrogen peroxide pre-treatment did not

change recovery of rhizobia significantly (Table 1) but reduced back

ground fluoresence, making rhizobia easier to count from non-rhizo

sphere soil. Cross-reactive bacteria were not a problem, and

cross-reactive fungal spores and mycelia were readily distinguishable

by their morphology. The soil contained no indigenous R. japonicum

capable of nodulating Davis soybeans. Nodules were observed on peas

tested with Kula soil, although very infrequently. These nodules were

small and white throughout, and bacteroids contained therein did not

react with FAs prepared for the strains used in this study.

Immunofluorescent examination of root surfaces indicated that the

majority of Rhizobium cells bound to roots were released into suspen

sion for counting.

Rhizosphere numbers of each of the four Rhizobium strains are

shown within each harvest across the three rhizospheres and fallow

soil (Figs. 1 to 4). Each point represents the mean of six enumerations

(duplicate filters from each of three replicate pots). Acridine orange

total counts for harvests 2 to 4 are presented in Figure 5. Each point

represents the mean of nine enumerations (triplicate filters from each

of three replicate pots).

Soybean nodules contained either of the two R. j aponicum strains

or both as illustrated in Figure 6. Pea nodules contained either of

30

Table 1. Recovery of added Rhizobium cells by the Kingsley and Bohlool method (11) and by a modification thereof*

Recovery method and strain

Cells recovered per gram of soil

Percentrecovery

Kingsley and Bohlool with flocculation of soil particles

USDA 110 4.5 X 1 0 6 70USDA 110 4.7 X 1 0 6 73USDA 110 4.7 X 1 0 6 73

Hawaii 5-0 4.2 X 106 77Hawaii 5-0 5.2 X 1 0 6 95Hawaii 5-0 5.0 X 1 0 6 92

Modified Kingsley and Bohlool with H2O2 pretreatment and centrifugation of soil particles

USDA 110 5.2 X 106 81USDA 110 5.1 X 106 80USDA 110 4.5 X 106 70

Hawaii 5-0 4.1 X 106 74Hawaii 5-0 4.4 X 106 81Hawaii 5-0 4.6 X 106 84

. *Average starting numbers of cells were USDA 110 6.4 x 106 and Hawaii 5-0 5.5 x 106. Recoveries were not significantly different (p=.05) by a Wilcoxon's signed rank test.

31

L.S.D, .05

FIG. 1 COUNTS OF 4 RHIZOBIUM STRAINS (X 106) PER GRAM OF OVEN DRY SOIL

IN 3 RHIZOSPHERES AND IN FALLOW SOIL AT HARVEST 1

32

L.S.D. .05

STRAIN

USDA 110

R. japonicum

CB 1809

N1TAAB1N 92A3

R. leguminosarum

HAHA1I 5-0

5.2THTBmr TEH CHHR

CHOPFHEcmr

FIG. 2 COUNTS OF 4 RHIZOBIUM STRAINS (X 10*) PER GRAM OF OVEN DRY SOIL


33

L • S . D . .05

HITRACIN 9203

R. leguminosarum

HflHflll 5-0

.05

18.0SOYBEAN

CROP



34

L • S - D • 05

NITRA61N 32R3

R. leguminosarum

HflHfllI 5-0

10.6

.05

FIG. 4 COUNTS OF 4 RHIZOBIUM STRAINS (X ltf) PER GRAM OF OVEN DRY SOIL


35

L .S .D. 05

HARVEST

FIG. 5 ACRIDINE ORANGE COUNTS OF BACTERIA IN 3 RHIZOPSHERES AND IN FALLOW SOIL (X 108)

PER GRAM OF OVEN DRY SOIL

36

HARVEST

10 20 30 40 50 80 70 80 90 tOOPERCENT

LEGENDi STRAINS SSR3S3 CB 1809 KSSSa MIXED V/////A USDA 110

FIG. 6 SOYBEAN NODULE OCCUPANCY BY RHIZOBIUM JAPONICUM STRAINS

the two inoculum R. leguminosarum strains or both as given in Figure 7.

Nodules were distributed throughout the root systems and were not

concentrated at the root crown. Mixed infections in nodules occurred

at an average of 15% for soybeans and 3.5% for peas. Of interest is

the occurrence of nodules on peas containing bacteroids unreactive

with either fluorescent antibody (Fig. 7). R. j aponicum bacteroids

were morphologically similar to broth cultured cells, whereas R.

leguminosarum bacteroids were enlarged relative to broth cultured

cells, and often branched (Figs. 8 and 9).

37

38

HARVEST

...I..""I.... I... "I.... I... ».|M ■■■ I ...p10 20 30 40 50 SO 70 80 90 100

PERCENTLEGEND i STRRINS m a m a HANA 11 5-0 1 U U J NIXED

B2ZZ3 NITRAGIN 92A3 ' i UNIDENTIFIED

FIG. 7 PEA NODULE OCCUPANCY BY RHIZOBIUM LEGUMINOSARUM STRAINS

39

FIG. 8 R. JAPONICUM NODULE BACTERIA, STAINED WITH HOMOLOGOUS FLUORESCENT ANTIBODY. CELLS ARE MORPHOLOGICALLY

SIMILAR TO BROTH CULTURED CELLS.(Scale = 10 ym)

FIG. 9 R. LEGUMINOSARUM NODULE BACTERIA, STAINED WITH HOMOLOGOUS FLUORESCENT ANTIBODY. CELLS ARE ENLARGED RELATIVE TO

BROTH CULTURED CELLS, AND OFTEN BRANCHED.(Scale = 10 ym)

DISCUSSION

One of the factors suggested as an important contributor to

specificity in the Rhizobium-legume association is the selective

stimulation of the homologous Rhizobium species in the rhizosphere of

its homologous host (7,14,25).

However, in this study I found that the growth of R. japonicum

was not selectively stimulated in the soybean rhizosphere, and that

the growth of R. leguminosarum was not selectively stimulated in the

pea rhizosphere. With the exception of harvest 2, rhizosphere numbers

of all strains of rhizobia were in the following order: soybeans>

peas>corn>non-rhizosphere soil, although differences in rhizosphere

populations of the crops are not always significant (Figs. 1 to 4).

During growth cycle 2 the plants were subjected to a considerable

short-term heat stress, due to the unfortunate failure of the air

conditioning unit on the plexiglas house. At harvest 2, rhizosphere

populations of the four Rhizobium strains were different than at

harvests 1, 3, and 4 (Figs. 1 to 4). Exudation from pea roots might

have been enhanced due to the heat stress. Soybean rhizosphere

populations of Rhizobium were reduced at harvest 2 relative to the

other harvests (Figs. 1 to 4). Apparently exudation from soybean

roots was reduced by the heat stress, or an inhibitory substance was

exuded. Hawaii 5-0 and USDA 110 were stimulated significantly more

in the rhizosphere of peas than in other rhizospheres at harvest 2

(Fig. 2). CB 1809 and Nitragin 92A3 were stimulated to the same

extent in the rhizospheres of peas and soybeans at harvest 2 (Fig. 2).

R. japonicum strains CB 1809 and USDA 110 were present in similar

numbers in any particular rhizosphere at any harvest (Figs. 1 to 4).

These strains apparently had similar rhizosphere colonizing abilities.

Hawaii 5-0 was generally seen in significantly lower numbers than the

other three strains in all rhizospheres at each of the four harvests

(Figs. 1 to 4). Hawaii 5-0 was apparently not as capable of rhizo

sphere colonization as the other strains in the rhizospheres tested.

In non-rhizosphere soil, CB 1809 was consistently seen in higher

numbers than the other strains (Figs. 1 to 4). Because this phenomenon

persisted through four harvests, it appeared that CB 1809 was better

adapted than the other strains to root-free soil.

Populations of individual Rhizobium strains never rose above

about 2 - 3 x 107 per gram of rhizosphere soil, even when soil was

continuously cropped with the same plant species. Thus, it seems as

though the rhizosphere might have a limited carrying capacity for

Rhizobium.

Bacteria enumerated with acridine organge were significantly more

prevalent in the soybean rhizosphere than the pea rhizosphere, than

the corn rhizosphere at harvests 3 and 4 (Fig. 5). At harvest 2,

total bacterial numbers were higher in the pea rhizosphere than the

soybean rhizosphere, probably due to the heat stress mentioned

earlier (Fig. 5). This pattern of rhizosphere colonization reflects

that seen with rhizobia, with bacteria more prevalent in the soybean

rhizosphere, than the pea rhizosphere, than the corn rhizosphere, with

the exception of harvest 2 (Figs. 2 to 4). Thus, the stimulation of

Rhizobium reflects the stimulation of rhizosphere bacteria in general.

42

The soybean nodule occupancy data were consistent with soybean

rhizosphere counts of the two R.. japonicum strains, as neither strain

predominated (Figs. 1 to 4 and 6). In the pea rhizosphere, Nitragin 92A3

was seen in numbers averaging twice those of Hawaii 5-0 (Figs. 1 to 4).

However, 92A3 was found in nine times as many nodules as Hawaii 5-0

(Fig. 7). Nitragin 92A3 appeared to be a more competent rhizosphere

colonizer than Hawaii 5-0, but increased rhizosphere populations of

Nitragin 92A3 do not fully explain the competitive advantage of this

strain in nodulating peas.

Hawaii 5-0 has been shown to be a very effective competitor in

nodulating lentils grown in several tropical soils, including an

Inceptisol (S. N. May, M.S. Thesis, 1979, University of Hawaii).

Perhaps the competitive ability of strains varies with respect to

different host species or cultivars, or with different soil types.

From these data, it appeared as though specific stimulation in

the rhizosphere was not a contributor to specificity in the R.

japonicum/soybean and R. leguminosarum/pea associations under the

conditions of this study. Differences in the growth of rhizobia in

host and non-host rhizospheres were manifested at the strain level

rather than at the species level. Thus some strains might simply be

better rhizosphere colonizers than others, without regard for

bacterial or plant species.

43

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46

25. Vincent, J. M. 1974. Root nodule symbiosis with Rhizobium.pp. 265-341. In A. Quispel (Ed.) The biology of nitrogen fixation. North Holland Publishing Co., Amsterdam.

POPULATIONS OF RHIZOBIUM IN THE RHIZOSPHERES OF HOST

AND NON-HOST PLANTS DURING A 35-DAY GROWTH CYCLE

CHAPTER 3

ABSTRACT

The growth of two R. j aponicum strains (USDA 110 and CB 1809) and

two R. leguminosarum strains (Hawaii 5-0 and Nitragin 92A3) was fol

lowed in the rhizospheres of soybean, pea, and corn plants growing in

non-sterile soil. Rhizosphere soil was sampled at weekly intervals

over a 35-day growth period. Rhizosphere numbers of each strain were

determined by membrane filter immunofluorescence, using strain-

specific fluorescent antibodies. Numbers of the four strains were

highest in the soybean rhizosphere at 1 week, averaging 2.0 x 107 per

gram of soil. Numbers of the four strains peaked in the pea rhizo-

sphere at 4 weeks, averaging 1.5 x 10 per gram of soil. At 1, 2, and

3 weeks, numbers per gram of soil of all strains were significantly

higher in the soybean rhizosphere than in the pea rhizosphere, than in

the corn rhizosphere, than in fallow soil. At week 4, pea rhizosphere

numbers per gram of soil of all strains significantly surpassed soybeans

rhizosphere numbers (1.5 x 107 and 1.2 x 107, respectively). At 5

weeks, rhizosphere numbers of all strains were in the following order:

pea>corn>soybean>fallow soil. Differences between Rhizobium numbers

in the rhizospheres of the three crops were more difficult to detect

on a root surface area basis. Numbers of all strains declined at 5

weeks per gram of soil and per square cm of root. No specific

stimulation of rhizobia by their legume hosts was observed. Strains

Nitragin 92A3, CB 1809, and USDA 110 were more successful rhizosphere

colonizers than Hawaii 5-0. Acridine orange total counts of bacteria

were highest on the basis of soil weight in the rhizospheres of the

three crops at 1 week, and following a decline at 2 weeks, remained

relatively constant for the remainder of the experiment. Acridine

orange total counts on the basis of root surface area were variable,

and patterns were not evident. Percentages of soybean nodules

occupied by the two R. j aponicum strains reflected the rhizosphere

populations of the strains, as neither predominated. Mixed

infections were observed at an average of 7%. The majority of large

nodules on peas contained Nitragin 92A3, and a lesser number,

Hawaii 5-0. Sixteen percent of large pea nodules contained R.

leguminosarum bacteroids unreactive with fluorescent antibodies

prepared against the two inoculant strains. In small pea nodules,

either one or both of the two inoculant strains were present in most

nodules at harvests 3 and 4. An unidentified strain or strains

occupied 74% of the small pea nodules at harvest 5. Mixed infections

in pea nodules varied from 4 to 10% in small and large nodules,

respectively. I found no evidence to support the theory of specific

stimulation of rhizobia by their homologous legume hosts in the R.

j aponicum/soybean and R. leguminosarum/pea associations.

49

INTRODUCTION

Rhizobia are distinguished from other soil bacteria by their

unique ability to form nitrogen-fixing nodules on the roots of many

leguminous plants. Rhizobia are specific with respect to the hosts

they nodulate. Only certain species of Rhizobium are able to infect

and nodulate the roots of particular legumes. Although the relation

ship between rhizobia and their host legumes has been studied in

great detail, the basis for this specificity is not completely clear

(3,16).

Several possible mechanisms have been proposed to account for

specificity, involving both plant and bacterial components (3).

Cellular recognition between host and bacterium is considered by

some as a mechanism for specificity. Plant proteins called lectins

are thought to interact specifically with rhizobia at the root

surface (6,17).

Another mechanism proposed as a possible contributor to

specificity in the Rhizobium-legume symbiosis is the preferential

growth of the appropriate Rhizobium in the rhizosphere of its

homologous host plant over and above that of other non-homologous

rhizobia, and over other soil microorganisms (3,10,19).

Most studies of stimulation in the legume rhizosphere have been

carried out in aseptic systems, from which rhizobia can be conve

niently plated and counted (5,9,11). Extrapolation from aseptic

conditions to non-sterile field soil, in which the full range of

microbial interactions would be occurring, is unrealistic, and

probably erroneous.

In a study described earlier (Chapter 2) rhizosphere numbers of

four strains of Rhizobium were determined in homologous and non-

homologous legume rhizospheres over four successive 35-day growth

cycles. Rhizobium populations were assessed on the basis of soil

weight by membrane filter immunofluorescence with strain-specific

fluorescent antibodies (1,8,15). Specific stimulation of homologous

rhizobia by their legume hosts was not observed, and rhizosphere

colonization by rhizobia reflected more strain differences than

species differences. It is possible however, that specific stimulation

could have occurred earlier in the growth cycle, possibly just prior

to the onset of nodulation. If this was the case, an experiment in

which rhizosphere sampling and enumeration was done at 35 days would

probably overlook specific stimulation.

Rovira (14) suggested that microbial densities in the rhizo

sphere on a unit soil weight basis be used with caution, since

rhizosphere soil weights are affected by plant species, soil moisture,

soil structure, and handling of the root during recovery of rhizo

sphere soil. He suggested that rhizosphere numbers be expressed on

the basis of root weight. Reyes and Schmidt (13) suggested that an

even more conservative method would be to express microbial densities

on the basis of root surface area, as roots of various plant species

vary in size, type and growth characteristics. They conducted a

series of experiments (12,13) to determine if stimulation of R.

japonicum strain 123 in the soybean rhizosphere is a possible

contributor to this strain’s extraordinary success in nodulating

soybeans in soils of the Midwest (7). They were unable to detect

any specific stimulation of strain 123 in the soybean rhizosphere.

51

What follows is the description of an experiment in which

membrane filter immunofluorescence was used to assess the population

dynamics of homologous and non-homologous rhizobia in the rhizospheres

of host and non-host plants over one 35-day growth cycle, with har

vests at weekly intervals. Immunofluorescence counts of rhizobia and

acridine orange counts of total bacteria are presented on the basis

of root surface area as well as rhizosphere soil weight.

52


Rhizobium Strains and Inoculum Preparation

Kula loam soil, an Inceptisol, was prepared as described

previously (Chapter 2). R. j aponicum strains CB 1809 and USDA 110,

and R. leguminosarum strains Hawaii 5-0 and Nitragin 92A3 were

obtained from the collection of B. B. Bohlool of the University of

Hawaii. Strains were maintained on yeast-extract mannitol agar

slants (18) with 1.0 g of yeast extract (Difco Laboratories, Detroit,

Michigan) substituted for yeast-water. Inoculum was prepared as

previously described (Chapter 2). Initial levels of the four

Rhizobium strains per gram of moist soil were as follows as determined

by membrane filter immunofluorescence: R. japonicum CB 1809,

4.7 x 105, and USDA 110 4.6 x 105, R. leguminosarum Hawaii 5-0,

3.8 x 105 and Nitragin 92A3 4.3 x 105.

Pots and Planting

Pots were 25 cm in diameter and 20 cm high and were painted with

a heavy coat of flat white paint. A watering device was placed into

each pot to provide water to the plants in the cores as evenly as

possible (Diagram 1). The pots were than each filled with 5.5 kg of

moist Kula soil. Cores of p.v.c. pipe of various sizes were then

driven into the soil in the pattern outlined in Diagram 2. The soil

inside of the cores was loosened slightly with a stout wire. The

soil was adjusted to about -1/3 bars of moisture tension with

deionized water. Three-day-old soybean, pea, and corn seeds (pre

germinated as described before) were planted, one per core. Following

Diagram I. View of one pot, with watering device installed.

20 m m long piece of 3/4" p.v.c. pipe, (notched at the bottom)

p.v.c. core

soil surface (covered with white aquarium gravel)

crushed gravel to a depth of 25 m m 150 x 25 m m plastic

petri plate bottom (Falcon Plastics 1013) Ui

55

Diagram 2. Arrangement of cores and order of harvest for the 35 - day time course study.

Perimeter of the watering device

Legend1. V'dia x 4" long p.v.c. pipe, 4. IV2" dia x 5" long p.v.c. pipe,harvested at 1 week harvested at 4" weeks.

2. 1" dia x 5"long p.v.c. pipe, 5. 2" dia x 5" long p.v.c. pipe,harvested at 2 weeks. harvested at 2 weeks.

3. 1" dia x 5" long p.v.c. pipe, harvested at 3 weeks.

planting, the soil surface was covered with rinsed white aquarium

gravel, to prevent undue soil heating and to inhibit the growth of

algae.

Plant Growth Conditions

The soil moisture tension was maintained at about -1/3 bars

throughout the experiment by watering to a constant weight with

deionized water. Water was poured into the central upright tube of

the watering device. In this manner, water was supplied to the roots

in the cores as evenly as possible from the perimeter of the base of

the device, rather than from a single central location. The experi

ment was set up as a randomized complete block with three replications

and carried out under natural lighting in a greenhouse.


At harvests 1, 2, and 3, four cores were removed from each pot

as outlined in Diagram 2. At harvests 4 and 5, two cores were

removed from each pot (Diagram 2). Root systems were separated from

the cores by gently tapping the cores on the inside of a metal pan.

Soil that adhered to the root system after gentle shaking was

considered to be rhizosphere soil. All other soil was returned to

the cores in the pots and gently compacted. Non-rhizosphere soil was

removed from the cores as above, pooled, mixed, and a 10-gram sample

taken. The remainder was placed back into the cores as described

above.

Bacteria were released from root surfaces and rhizosphere soil

particles as previously described (Chapter 2). At harvest 1, 50 ml

56

of the gelatin-ammonium phosphate mixture (8) was used for extractions

rather than 100 ml, because of the small amount of rhizosphere soil

present.

Root Surface Area Estimation

Root surface area was estimated by the method of Carlson (4), as

modified by Reyes and Schmidt (13) for greenhouse-grown soybeans.

Enumeration of Bacteria

Membrane filter counts were performed as described by Kingsley

and Bohlool (8). Duplicate filters were counted for each strain from

each pot, for a total of 96 filters per harvest.

Acridine orange total counts were made at each harvest as

previously described. Three filters were counted per pot, for a

total of 36 per harvest.

Nodule Typing

All nodules were removed, squashed, smeared on microscope

slides, and serologically typed with fluorescent antibodies as

previously described (Chapter 2).

57

RESULTS

Rhizosphere numbers of each of the four inoculant strains are

presented at weekly intervals on the basis of soil weight in Figures 1

to 5. Numbers of all strains per gram of rhizosphere soil are in the

following order at 1, 2, and 3 weeks: soybean>pea>corn>non-rhizosphere

soil (Figs. 1 to 3). At 4 weeks rhizobia per gram of soil had declined

in the soybean rhizosphere, but increased in that of peas, signifi

cantly surpassing soybean rhizosphere numbers (Fig. 4). Rhizobium

numbers per gram of soil of all strains were higher in the soybean

rhizosphere than that of corn at 4 weeks, with the exception of

Hawaii 5-0 (Fig. 4). At 5 weeks, numbers of rhizobia per gram of soil

were in the following order: pea>corn>soybean>non-rhizosphere soil.

Rhizosphere numbers of each of the four strains are presented at

weekly intervals on the basis of root surface area in Figures 6 to 10.

Differences in rhizosphere populations of Rhizobium between the crops

were more difficult to detect on the basis of root surface area,

owing to the variability in the estimations of root surface area

(Figs. 6 to 10).

Rhizosphere bacteria enumerated using acridine orange fluores

cence microscopy were in the following order: soybean>pea>corn>non-

rhizosphere soil. However, the variability of the acridine orange

total counts was high, and differences between crops and non-rhizo

sphere soil are often not significant (Fig. 11). Differences between

rhizosphere populations of bacteria per square cm of root between

crops were even more difficult to detect, with the only significant

differences at 5 weeks (Fig. 12).

59



* denotes L.S.D. of less than 0.05

60

L.S.D. .05

0 .0* 0 .0*

STRAIN

R. japonicum

NITRAGIN

R. leguminosarum

HANAII 5-0



.2

' .05


61

L.S.D. .05

n . s. 0.3 0.1 0 .0*

CROP

USDR 110

R. japonicum

CB 1808

HflHflll 5-0

N1TRRGIN 92R3

R. leguminosarum




62

L.S.D. .05

N1TRA81N 9203

R. leguminosarum

HflHflll 5-0

STRAIN

USDA 110

R. iaponicum

CB 1808

L.S.D

FALLONCROP




63

L.S.D. .05

n.s. 0.1 0.1

L.S.D

STRAIN

USDA 110

R. japonicum

CB 1809

NITnASIN 92B3

R. leguminosarumHAMAI1 5-0



64

L.S.D..05

1.4 0.3

/I / n.s.

n. s.

L.S.D. .05

n.s.

FIG. 6 COUNTS OF 4 RHIZOBIUM STRAINS (X 10*) PER SQUARE CENTIMETER OF ROOT AREA

IN 3 RHIZOSPHERES AT HARVEST 1

65

L.S.D. 05

CROP

L.S.D.

U3DR 110

R. japonicum

CB 1B09

NITRAGIN 92A3

R. leguminosarum

HAWAII 5-0

U. 7

FIG. 7 COUNTS OF 4 RHIZOBIUM STRAINS (X 104) PER SQUARE CENTIMETER OF ROOT AREA


66

L.S.D. .05

n.s.

US0A 110

R. japonicum

CB 1809

NITRAGIN 92R3

CROP

n.s. L.S.D.

n.s.R. leguminosarum

HflNRlI 5-0

FIG. 8 COUNTS OF 4 RHIZOBIUM STRAINS (X 104) PER SQUARE CENTIMETER OF ROOT AREA


S .

.05

67

FIG. 9 COUNTS OF 4 RHIZOBIUM STRAINS (X 10A) PER SQUARE CENTIMETER OF ROOT AREA


STRAIN

n.s. 0.3

1.3L.S.D.

NITRAGIN 82A3

R. leguminosarum

CROP

USOA 110

R. japonicum

CB 1808

12.4

12.6

FIG. 10 COUNTS OF 4 RHIZOBIUM STRAINS (X 10A) PER SQUARE CENTIMETER OF ROOT AREA


69

HAHVE3T

FIG. 11 ACRIDINE ORANGE TOTAL COUNTS OF BACTERIA (X 108) PER GRAM OF OVEN DRY SOIL

IN 3 RHIZOSPHERES AND IN FALLOW SOIL

HARVEST

FIG. 12 ACRIDINE ORANGE TOTAL COUNTS OF BACTERIA (X 107) PER SQUARE CENTIMETER OF ROOT AREA

IN 3 RHIZOSPHERES

Soybean nodules contained the two inoculant R. japonicum strains

as outlined in Figure 13. Mixed infections were observed at an

average of 7%. Large and small nodules formed on pea plants. Large

nodules were elongated and red inside. Small nodules were nearly

spherical, and white throughout. Large and small nodules contained

the two inoculant R. 1eguminosarum strains as well as an unidentified

strain or strains as outlined in Figures 14 and 15. Mixed infections

occurred in large and small pea nodules at an average of 10 and 5%,

respectively.

Photographs of the experiment are presented at 1, 3, and 5 weeks

in Figures 16, 17, and 18.

71

72

HARVEST

1 2

3

4

510 20 30 40 50 60 70 80 90 100

PERCENTLEGENO> STRAINS ESSES CB 1809 B C T MIXED USDA 110

FIG. 13 SOYBEAN NODULE OCCUPANCY BY RHIZOBIUM JAPONICUM STRAINS

73

HARVEST

2

3

4

5

- - - - - -

8 §10 20 30 40 50 60 70 80 90 100

PERCENTLEGENDi STRAINS l w w a i HAWAII 5-0

NITRAGIN 92A3LAAAAJ m i x e d i 1 UNIDENTIFIED

FIG. 14 LARGE PEA NODULE OCCUPANCY BY RHIZOBIUM LEGUMINOSARUM STRAINS

74

HARVEST

3

4

5

tO 20 30 40 50 80 70 80 90 tOOPERCENT

LEGENDt STRAINS HAWAII 5-0 lxaaaj MIXEDZZZZZZZ NITRAGIN 92A3 i 1 UNIDENTIFIED

FIG. 15 SMALL PEA NODULE OCCUPANCY BY RHIZOBIUM LEGUMINOSARUM STRAINS

. 16 VIEW OF THE EXPERIMENT AT 1 WEEK. GROWTH OF THE EXPERIMENTAL PLANTS WAS UNIFORM WITHIN EACH SPECIES,

AND WITHIN AND BETWEEN REPLICATE POTS.

77

FIG. 17 VIEW OF THE EXPERIMENT AT 3 WEEKS. PLANTS GREW WELL DESPITE THE CONFINEMENT OF THEIR ROOT SYSTEMS BY CORES.CORN PLANTS ARE STARTING TO SHOW NITROGEN DEFIEIENCY

SYMPTOMS (CHLOROSIS OF OLDER GROWTH), WHEREAS THE LEGUMES ARE GREEN AND HEALTHY.

FIG. 18 VIEW OF THE EXPERIMENT AT 5 WEEKS. AT 5 WEEKS, PEAS WERE FILLING PODS AND SOYBEANS HAD FLOWERED. THE NITROGEN

DEFICIENCY OF CORN WAS MORE PRONOUNCED, AS OLDER LEAVES WERE SEVERELY CHLOROTIC OR NECROTIC.

DISCUSSION

One of the factors proposed as a possible contributor to

specificity in the Rhizobium-legume symbiosis is the selective

stimulation of the homologous Rhizobium species in the rhizosphere of

its homologous host (3,10). In the previous study (Chapter 2) I

found that species of rhizobia were not specifically stimulated in

the rhizospheres of their host at 35 days. I designed this study to

examine the population dynamics of rhizobia in host and non-host

rhizospheres at weekly intervals during the growth of the plants.

Reyes and Schmidt (13) suggested that root surface area might be a

better basis for expressing rhizosphere populations of rhizobia, so

root area was estimated, and data presented on the basis of root area

as well as soil weight.

As was seen previously (Chapter 2), rhizosphere stimulation of

host and non-host rhizospheres was dependent more on the Rhizobium

strains than on the particular species of Rhizobium. At 1, 2, and

3 weeks, numbers per gram of soil of the strains were in the follow

ing order: soybean rhizosphere>pea>corn>non-rhizosphere soil

(Figs. 1 to 3). Differences between crops at these harvests were

all significant at the .05 level. At harvests 4 and 5 soybean

rhizosphere* numbers decreased (Figs. 4 and 5). At harvest 5 both pea

and corn rhizospheres harbored significantly more rhizobia of all

strains than the soybean rhizosphere (Fig. 5). This sharp decline

of all strains in the soybean rhizosphere might be an artifact of

having grown the plants in cores. At 4 and 5 weeks much of the

soybean root system had emerged from the bottom of the cores, and

roots proliferated rapidly in the bottom of the pots. The lowered

numbers in the rhizospheres of peas and soybeans at 5 weeks probably

reflect the fact that roots were growing rapidly, and enmeshing

rhizosphere soil faster than the growth of the inoculant strains

could keep up.

Differences in rhizosphere populations of rhizobia in the rhizo

spheres of the three crops were harder to detect on the basis of root

surface area. Patterns of rhizosphere colonization were similar to

those observed when rhizosphere populations were examined on a soil

weight basis (Figs. 6 to 10). Significant differences between crops

were only observed at 2 and 5 weeks (Figs. 7 and 10). Variability

between root surface area estimations was high. This variability

precluded the detection of differences between Rhizobium numbers in

the rhizospheres of the three crops for harvests 1, 3, and 4 (Figs. 6,

8, and 9). Significant differences were observed between strains on

the basis of root surface area across the five harvests, however

differences were not indicative of specific stimulation (Figs. 6 to

10). As was seen previously (Chapter 2), numbers of the two R.

japonicum strains were generally similar, and similar to numbers of

R. leguminosarum strain Nitragin 92A3. R. leguminosarum strain

Hawaii 5-0, however, is seen in significantly lower numbers than the

other three strains in all rhizospheres and across all five harvests

(Figs. 1 to 10). This would indicate rhizosphere colonization more

on the basis of differences between strains, rather than species

differences, with Hawaii 5-0 being less adapted to the conditions in

the rhizospheres of all plants.

80

As was seen previously (Chapter 2), soybean nodule occupancy data

approximated the rhizosphere counts of USDA 110 and CB 1809, as

neither dominated (Fig. 13). In large pea nodules, Nitragin 92A3

dominated Hawaii 5-0 with the exception of harvest 2, when percentages

of the two strains were nearly equal (Fig. 14). Nodule occupancy of

small pea nodules was different. At harvest 3, both inoculant strains

occupied nodules in approximately equal percentages, and unidentified

bacteroids occupied a nearly equal percentage (Fig. 15). However,

Hawaii 5-0 occupied more small nodules than Nitragin 92A3 at harvests

4 and 5, and at harvest 5 unidentified bacteroids predominated

(Fig. 15).

Hawaii 5-0 has been shown to be very competitive in nodulating

lentils in many tropical soils, including an Inceptisol (S. N. May,

M.S. Thesis, 1979, University of Hawaii). However, Hawaii 5-0 was

not as competitive as Nitragin 92A3 in nodulating peas. Perhaps the

soil or host plant influence the competition between strains for

nodulation of legumes.

Plants grew well in the Kula soil, despite having their root

systems confined to cores. Lack of uniformity within or between

replicates was not a problem (Figs, 16, 17, and 18). At 5 weeks, the

corn was slightly chlorotic relative to the legumes, probably due to

a nitrogen deficiency, as no amendments (other than rhizobia) were

made to the soil (Fig. 18).

In this study, no evidence was found in support of specific

stimulation of R. j aponicum in the rhizosphere of soybeans or R.

1eguminosarurn in the rhizosphere of peas. Rhizosphere colonization

81

by rhizobia reflected more strain differences than species

differences.

Rhizosphere numbers of rhizobia were quite high in the soybean

and pea rhizospheres at 1 week, increasing from about 4 - 5 x 105 per

gram of moist soil at zero time to 1 - 2 x 107 per gram of oven dry

soil at 1 week. Although no specific responses were observed on the

basis of rhizosphere growth during this study, it is possible that

specific interactions occurred before the first sampling time, at

1 week. This possibility will be investigated in a later experiment.

82

LITERATURE CITED

Bohlool, B. B. and E. L. Schmidt. 1973. A fluorescent antibodytechnique for determination of growth rates of bacteria in soil.Bull. Ecol. Res. Comm. (Stockholm) 17:229-236.

Broughton, W. J. 1978. Control of specificity in legume-Rhizobium associations. J. Appl. Bacteriol. 45:165-194.

Brown, Margaret E., R. M. Jackson, and S. K. Burlingham. 1968. Growth and effects of bacteria introduced into soil. In T. R. G. Gray and D. Parkinson (Eds.) The ecology of soil bacteria. Liverpool Univ. Press, Liverpool, pp. 531-551.

Carlson, J. B. 1969. Estimating surface area of soybean root systems. J. Minn. Acad. Sci. 36:16-19.

Van Egeraat, A. W. S. M. 1975. The growth of Rhizobiumleguminosarum on the root surface and in the rhizosphere in relation to root exudates. Plant and Soil 42:367-379.

Graham, Terrence L. 1981. Recognition in Rhizobium symbioses.In Kenneth L. Giles and Alan G. Atherly (Eds.) Biology of the Rhizobiaceae. Academic Press, Inc., New York.

Ham, G. E., L. R. Frederick, and L. C. Anderson. 1971. Serogroups of Rhizobium j aponicum in soybean nodules sampled in Iowa. Agron.J. 63:69-72.

Kingsley, Mark T. and B. B. Bohlool. 1981. Release of Rhizobium spp. from tropical soils and recovery for immunofluorescence enumeration. Appl. Environ. Micro. 42:241-248.

MacGregor, A. N. and M. Alexander. 1972. Comparison of nodulating and non-nodulating strains of Rhizobium trifolii. Plant and Soil 36:129-139.

Nutman, P. S. 1965. The relation between nodule bacteria in the rhizosphere and in the process of infection. In K. F. Baker and W. C. Snyder (Eds.) The ecology of soil-borne plant pathogens, pp. 231-247. Univ. Calif. Press, Berkeley.

Peters, R. J. and M. Alexander. 1966. Effect of legume exudates on the root nodule bacteria. Soil Sci. 102:380-387.

Reyes, V. G. and E. L. Schmidt. 1979. Population densities of Rhizobium japonicum strain 123 estimated directly in soil and rhizospheres. Appl. Environ. Micro. 37:854-858.

84

13. Reyes, V. G. and E. L. Schmidt. 1981. Populations of Rhizobiumj aponicum associated with the surfaces of soil-grown roots. Plant and Soil 61:71-80.

14. Rovira, A. D. 1961. Rhizobium numbers in the rhizosphere of redclover and paspalum in relation to soil treatment and numbers ofbacteria and fungi. Aust. J. Agric. Res. 12:77-83.

15. Schmidt, E. L. 1974. Quantitative autoecological study of microorganisms in soil by immunofluorescence. Soil Sci. 118:141-149.

16. Schmidt, E. L. 1978. Ecology of the legume root nodule bacteria,pp. 269-303. In Y. R. Dommergues and S. V. Krupa (Eds.) Interactions between non-pathogenic soil microorganisms and plants. Elsevier Scientific Publishing Co., Amsterdam.

17. Schmidt, E. L. and B. B. Bohlool. 1981. The role of lectins inplant-microbe interactions, pp. 658-677. In W. Tanner andF. A. Loewus (Eds.) Encyclopedia of plant physiology, New Series, Vol. 4. Plant carbohydrates II. Springer-Verlag, Berlin, Heidelberg.

18. Vincent, J. M. 1970. A manual for the practical study of rootnodule bacteria. International Biological Programme Handbook no. 15. Blackwell Scientific Publications, Oxford, England.

19. Vincent, J. M. 1974. Root nodule symbiosis with Rhizobium.pp. 265-341. In Quispel (Ed.) The biology of nitrogen fixation. North Holland Publishing Co., Amsterdam.

RHIZOBIUM POPULATION DYNAMICS IN HOST AND NON-HOST RHIZOSPHERES

DURING THE FIRST NINE DAYS OF RHIZOSPHERE DEVELOPMENT

CHAPTER 4

ABSTRACT

The growth of two R. japonicum strains (USDA 110 and CB 1809) and

two R. leguminosarum strains (Hawaii 5-0 and Nitragin 92A3) was fol

lowed in non-sterile rhizospheres. In one study, growth of these

strains was followed in the rhizospheres of soybeans and peas at

planting, 24, and 48 hours, and rhizosphere growth rates estimated.

Mean doubling times for the strains were 12 to 13 hours in the soybean

rhizosphere and 9 to 11 hours in the pea rhizosphere. Total counts of

bacteria measured by acridine orange staining indicated that other

bacteria were not as strongly stimulated by the rhizosphere as

rhizobia in the early stages of plant growth. In another study,

rhizosphere populations of the four strains were followed in the

rhizospheres of soybeans, peas, and corn, as well as in non-rhizo

sphere soil at 3, 5, 7, and 9 days. Rhizosphere numbers of Rhizobium

increased from their starting levels of 4 - 9 x 10s per gram of moist

soil to 1 - 2 x 107 per gram of moist soil at 3 days, and declined

thereafter. Populations of bacteria enumerated with acridine orange

remained relatively constant. This early burst of the growth of

rhizobia might indicate that they responded faster to the conditions

of the rhizosphere than the bacterial population at large, but were

diluted out as the root systems expanded rapidly. No specific stimu

lation of the growth of rhizobia was detected in the rhizospheres of

their host plants. Rhizosphere populations reflected strain differ

ences more than species differences, in all crops and at all harvests.

INTRODUCTION

Soil bacteria of the genus Rhizobium are distinguished from other

genera by their ability to form nitrogen-fixing nodules on the roots

of leguminous plants. Rhizobia are specific with respect to the hosts

they nodulate. Only certain species of Rhizobium are able to infect

and nodulate the roots of particular legumes. Although the relation

ship between host and bacterium has been studied in great detail, the

basis for this specificity is not completely clear (1,7).

One proposed mechanism as a possible contributor to specificity

is the preferential growth of the appropriate Rhizobium in the

rhizosphere of its homologous host plant over other soil bacteria and

rhizobia (2,4,9).

In studies described earlier (Chapters 2 and 3), rhizosphere

numbers of four Rhizobium strains were determined at harvest across

four 35-day harvests, as well as at weekly intervals during one 35-day

growth cycle. I observed no specific stimulation of growth of

Rhizobium species in the rhizospheres of their homologous host plants

in these studies. When rhizosphere populations were sampled once a

week, numbers of all strains increased from their starting levels of

about 4 x 105 per gram of soil to a level of about 1 x 107 per gram of

soil at 1 week, and generally declined thereafter.

Following is the description of two experiments in which population

dynamics of homologous and non-homologous rhizobia were followed in the

rhizospheres of host and non-host plants. Rhizosphere populations were

sampled in one study at planting, 24, and 48 hours, and in another

study at 3, 5, 7, and 9 days.


Inoculum Preparation and Rhizobium Strains

Fresh Kula loam soil, prepared as described previously, was used

for the 9-day time course study. Initial levels of each of the strains

per gram of moist soil were as follows, as determined by immuno

fluorescence: R. j aponicum USDA 110, 8.3 x 105 and CB 1809, 9.2 x 105,

_R. leguminosarum Nitragin 92A3, 4.2 and 105 and Hawaii 5-0, 3.9 x 105.

For the early rhizosphere growth rate study, about 5 x 109 cells of

each strain (as estimated by optical density) were transferred from

yeast-extract mannitol broth (8) culture and blended with 2000 grams

of fresh Kula soil, to give about 2.5 x 106 cells per gram of moist

soil. This intermediate dilution was blended with more moist Kula soil

275 g of intermediate dilution to 5225 g of bulk soil. This 1:20

dilution was expected to give about 1.25 x 105 cells per gram of moist

soil. Actual starting numbers at planting were slightly higher than

this estimate, as follows: USDA 110, 4.3 x 10s, CB 1809, 5.1 x 105,

Nitragin 92A3, 4.3 x 105 and Hawaii 5-0, 4.7 x 105. Strains were

obtained from the collection of B. B. Bohlool of the University of

Hawaii, and were grown and maintained as described previously.

Pots and Planting

Pots were 25 cm in diameter and 20 cm high, and were painted with

a heavy coat of flat white paint. Each was filled with 5.5 kg of

moist Kula soil. Into each pot was planted 80 seeds. Soybeans (Davis)

peas (Wilt-resistant Wisconsin Perfection), and sweetcorn (Hawaiian

Supersweet #9) were the crops. In the growth rate study, only peas and

soybeans were planted. Seeds were not pregerminated. Following

planting, the soil surface was covered with rinsed white aquarium

gravel to prevent undue soil heating and the growth of algae.

Deionized water was added to bring the soil to a moisture tension of

about -1/3 bars.

Soil moisture tension was maintained at about -1/3 bars by

watering to constant weight with deionized water. Each experiment was

set up as a randomized complete block in three replications and carried

out in a greenhouse under natural lighting conditions. Soil temper

ature during the afternoon was about 30°C.


For the 9-day time course, I harvested 15 to 20 plants from each

pot at 3, 5, 7, and 9 days after the planting of dry seeds. Forty

plants were dug out (by hand) from each pot at 24 and 48 hours in the

growth rate study. Rhizosphere soil was considered to be that which

adhered to seeds and root system after gentle shaking. Plants and

root systems were placed into 18-ounce Whirlpak bags for transport to

the laboratory. A sample of approximately 10 grams was removed from

the non-rhizosphere soil pots at each time point during the 9-day time

course. Ten-gram samples were removed from pea and soybean pots

immediately before the planting of the growth rate study.

Bacteria were released from root surfaces and rhizosphere soil

particles for enumeration as previously described (Chapter 2). At

harvest 1 of the 9-day time course, 50 ml of the extractant mixture

(3) was used rather than 100 ml, because of the small amount of rhizo

sphere soil present. Twenty and 50 ml of extractant were used for the

24-and 48-hour enumerations, respectively, of the growth rate study.

89

Membrane Filter Immunofluorescence and Acridine Orange Total Counts

90

Membrane filter counts were performed as described by Kingsley

and Bohlool (3). Duplicate filters were counted for each strain from

each pot, for a total of 96 filters per harvest in the 9-day time

course and 48 per harvest of the growth rate study.

I made acridine orange counts at each harvest as previously

described (Chapter 2). Three filters were counted per pot, for a

total of 36 per harvest in the time course and 18 per harvest in the

growth rate study.

RESULTS

Rhizosphere numbers of each of the four inoculant strains are

presented at 3, 5, 7, and 9 days in Figures 1 to 4. Acridine orange

total counts at the above time points are presented in Figure 5.

Populations of each of the four inoculant strains are presented

at planting, 24, and 48 hours in the pea and soybean rhizospheres in

Figures 6 and 7. Estimates of mean doubling times of inoculant strains

are also presented in Figures 6 and 7. At 24 hours, soybeans had

radicles about 1 cm long, and peas were swollen, but radicles had not

yet extended. At 48 hours, soybeans and peas had radicles of about

6 cm and 1 cm, respectively. Rhizosphere populations of bacteria

estimated by acridine orange total counts are presented at planting,

24, and 48 hours in Figure 8.

Percentages of Rhizobium (the four inoculant strains combined) in

the rhizosphere population of bacteria are given for each study in

Tables 1 and 2, respectively.

Nodules were not observed on the roots of the legumes, due to the

short duration of the experiment.

92

L.S.D. .05

n.s. n.s. 0.4

L.S.D

13.5

chop

USDA ITO

R. japonicumCB 1809

N1TIW61N 92H3

R. leguminosarumHAHA11 5-0


IN 3 RHIZOSPHERES AND IN FALLOW SOIL AT 3 DAYS

93

L.S.D. .05

CROP

L.S.D.

usoa 110

R. japonicumCB 1809

N1TIW91N 8203

R. leguminosarum

HflHflll 5-0



94

FIG. 3 COUNTS OF 4 REIZOBIUM STRAINS (X 105) PER GRAM OF OVEN DRY SOIL


95

L.S.D. .05

CROP

STRAIN

USDA 110

R. japonicumCB 1809

HAWAII 5-0

N1TRA61N 92A3

R. leguminosarum

L.S



96

HARVEST

FIG. 5 ACRIDINE ORANGE TOTAL COUNTS OF BACTERIA (X 108) PER GRAM OF OVEN DRY SOIL

IN 3 RHIZOSPHERES AND IN FALLOW SOIL

97


IN THE PEA RHIZOSPHERE AT PLANTING, 24, AND 48 HOURS

(10.0)

(8.9)

7)

(hours)

98


IN THE SOYBEAN RHIZOSPHERE AT PLANTING, 24, AND 48 HOURS

99

L.S.D. .05

FIG. 8 ACRIDINE ORANGE TOTAL COUNTS OF BACTERIA (X 108) PER GRAM OF OVEN DRY SOIL IN PEA AND SOYBEAN RHIZOSPHERES

Table 1. Percentages of identified Rhizobium in the rhizosphere population of bacteria at 3, 5,

7, and 9 days

Time from planting (days) 3 5 7 9

Rhizosphere

soybean 12 6 5 5

pea 12 10 5 7

corn 9 3 3 4

non-rhizosphere 5 4 4 4soil

101

Table 2. Percentages of identified Rhizobium in the rhizosphere population of bacteria at

planting, 24, and 48 hours

Time from planting (hours) 0 24 48

Rhizosphere

soybean 1.0 1.3 5.6

pea 0.8 2.0 8.0

DISCUSSION

Specific stimulation in the rhizosphere is proposed as a possible

contributor to specificity in the Rhizobium-legume symbiosis (2,4,9).

In two earlier studies (Chapters 2 and 3), population dynamics of two

R. japonicum and two R. leguminosarum strains was followed in host and

non-host rhizospheres. I sampled rhizosphere populations at 35 days

in one experiment and at weekly intervals during one 35-day growth

cycle in another. In these studies, as well as those of Reyes and

Schmidt (5,6) on the soybean R. japonicum strain 123 association, no

specific stimulation was detected in the rhizosphere.

Rhizosphere numbers of rhizobia were in the following order:

soybean>pea>corn at 3, 5, and 7 days, although differences between

rhizosphere populations between crops were not always significant

(Figs. 1 to 3). At 9 days, pea rhizosphere numbers of all strains

significantly surpassed those for soybean, with the exception of

strain Nitragin 92A3 (Fig. 4). Rhizobia per gram of soybean rhizo

sphere soil declined over the four harvests (Figs. 1 to 4). In the

pea rhizosphere rhizobia declined for the first week (Figs. 1 to 3),

then increased slightly at 9 days (Fig. 4). Numbers of Rhizobium per

gram of corn rhizosphere soil were rarely significantly different from

numbers in non-rhizosphere soil at 3, 5, 7, and 9 days. As was seen

in Chapters 2 and 3, numbers of the two R. japonicum strains were

nearly equal in any particular rhizosphere at any given harvest.

Rhizosphere numbers of Hawaii 5-0 were sometimes significantly lower

than numbers of the other three inoculant strains, but differences

were usually slight, less than the two-fold differences seen in

Chapter 2 (Figs. 1 to 4). Apparently the competitive advantage of the

other strains over Hawaii 5-0 was not as great during early development

of the rhizosphere.

In the growth rate study, rhizobia responded rapidly to the condi

tions of the rhizosphere, and numbers of each strain increased about

1 1/2 logs in the first 48 hours (Figs. 6 and 7). Growth rates in the

early rhizosphere were roughly equal for both the fast-growing (R.

leguminosarum) and the slow-growing (R. japonicum) strains. This

suggests that there might be limiting nutrients or growth factors

which restrict the growth of rhizobia, regardless of their growth rate

in broth culture.

The soybean rhizosphere did not specifically stimulate the growth

of R. j aponicum, nor did the pea rhizosphere specifically stimulate R.

leguminosarum (Figs. 1 to 4, 6 and 7). In the 9-day time course, as

was seen previously (Chapter 3) numbers of all inoculant strains of

rhizobia in all rhizospheres were highest at the first time point (3

and 7 days for the two studies, respectively). At 3 days rhizobia

comprised their highest percentages of the rhizosphere population of

bacteria, relative to acridine orange total counts (Tables 1 and 2).

It appears that rhizobia responded to the conditions of the early

rhizosphere with a burst of rapid growth. Rhizobia multiplied rapidly

in the developing seedling rhizosphere, and reached peak populations,

on the basis of soil weight, even before the emergence of shoots from

the soil. Thus, the early events in the rhizosphere may play an

important role in the nodulation of legumes, as proposed by Kosslak,

Dowdle, Sadowsky, and Bohlool (8th North American Rhizobium Conference,

103

1981. Abstracts). However, patterns of growth do not reflect

selective stimulation of rhizobia by their homologous hosts (Figs. 1 to

4, 6 and 7). Soybean and pea rhizospheres stimulated the growth of

rhizobia more strongly than the corn rhizosphere during the first nine

days of growth (Fig. 1). This was also the case for bacteria in

general (Fig. 5).

Rhizobial numbers declined after this initial burst of growth,

probably because the plant root systems expanded and enmeshed soil

faster than rhizobia could colonize it.

These data support the work of Reyes and Schmidt (5,6), who

examined the growth of R. japonicum strain 123, a highly competitive

soybean strain, both on the basis of rhizosphere soil weight (5) and on

the basis of root surface area (6). They found no evidence to support

the concept of specific stimulation as a possible contributor to

specificity. In the first study (5), rhizosphere numbers of strain 123

per gram of soil peaked at 21 days, and declined at 28 days. In the

second experiment (6), they observed a decline in numbers of strain 123

in the soybean rhizosphere between days 16 and 30, on the basis of root

area. Reyes and Schmidt (6) postulated that the rhizosphere population

of strain 123 is unable to keep up with the rapidly-expanding soybean

root system.

It is possible that specific stimulation might have occured at

microsites along the plant root system. If this were the case, the

rhizosphere sampling and enumeration methodology such as the one used

for these studies might be inadequate to detect the stimulation. This

highly, localized stimulation seems unlikely for the case of the soybean

rhizosphere, however. Reyes and Schmidt (6) examined soil adhering

104

to unwashed soybean roots and found populations of two strains of R.

j aponicum to be uniform and low (6).

Acridine orange total counts were relatively constant throughout

the 9-day time course (Fig. 5). An early burst of growth of bacteria

was observed in the first 48 hours. Rhizobia grew more rapidly than

the other soil bacteria, but because of the high initial numbers of

other bacteria, they always comprised the vast majority of the rhizo

sphere population (Tables 1 and 2). This early burst of growth was

not specific with respect to legume host or Rhizobium species.

However, this early burst of growth in the rhizosphere might be a

factor in ensuring the successful establishment of the Rhizobium-

legume symbiosis.

105

LITERATURE CITED

Broughton, W. J. 1978. Control of specificity in legume- Rhizobium associations. J. Appl. Bacteriol. 45:165-194.

Brown, Margaret E., R. M. Jackson, and S. K. Burlingham. 1968. Growth effects of bacteria introduced into soil. In T. R. G. Gray and D. Parkinson (Eds.) The ecology of soil bacteria. Liverpool Univ. Press, Liverpool, pp. 531-551.

Kingsley, Mark T. and B. B. Bohlool. 1981. Release of Rhizobium spp. from tropical soils and recovery for immunofluorescence enumeration. Appl. Environ. Micro 42:241-248.

Nutman, P. S. 1965. The relation between nodule bacteria in the rhizosphere and in the process of infection. In K F. Baker and W. C. Snyder (Eds.) The ecology of soil-borne plant pathogens, pp. 231-247. Univ. Calif. Press, Berkeley.

Reyes, V. G. and E. L. Schmidt. 1979. Population densities of Rhizobium japonicum strain 123 estimated directly in soil and rhizospheres. Appl. Environ. Micro. 37:854-858.

Reyes, V. G. and E. L. Schmidt. 1981. Populations Rhizobium japonicum associated with the surfaces of soil-grown roots. Plant and Soil 61:71-80.

Schmidt, E. L. 1978. Ecology of the legume root nodule bacteria,pp. 269-303. In Y. R. Dommergues and S. V. Krupa (Eds.) Interactions between non-pathogenic soil microorganisms and plants. Elsevier Scientific Publishing Co., Amsterdam.

Vincent, J. M. 1970. A manual for the practical study of theroot nodule bacteria. International Biological Programme Handbookno. 15. Blackwell Scientific Publications, Oxford, England.

Vincent, J. M. 1974. Root nodule symbiosis with Rhizobium. pp. 265-431. In Quispel (Ed.) The biology of nitrogen fixation. North Holland Publishing Co., Amsterdam.

CHAPTER 5

GENERAL DISCUSSION

GENERAL DISCUSSION

From these data it would appear that specific stimulation in the

rhizospheres of peas and soybeans of their homologous Rhizobium species

was not seen under the conditions of these studies. Bacteria, including

rhizobia, were generally more strongly stimulated by the soybean

rhizosphere than the pea rhizosphere. The rhizosphere of sweetcorn

harbored more bacteria than fallow soil, but less bacteria than the

legume rhizospheres.

All strains grew rapidly in the early periods of rhizosphere

development. Numbers peaked at about 3 days in the pea and soybean

rhizospheres, and declined thereafter as the plant root systems expanded

rapidly. Nodulation patterns reflected rhizosphere populations of the

Rhizobium strains, but rhizosphere differences alone could not account

for the differences in nodule occupancy.

Rhizosphere populations of Rhizobium reflected more strain differ

ences than species differences. Perhaps rhizobia colonize rhizospheres

without regard to plant species, and some strains are better rhizo

sphere colonizers than others.

Curiously, rhizosphere populations of each strain never got above

about 2 - 3 x 107 cells per gram of soil even when soil was continuously

cropped with one plant species. Thus, it seems as though the rhizosphere

might have a limited carrying capacity for Rhizobium.

The rapid early growth of Rhizobium in the rhizosphere was

surprising. Although the growth of all inoculant strains was similar in

the early rhizosphere in these experiments, this might not be the case

for all strains in all rhizospheres.

Further work should concentrate on the documentation of the rapid

growth of rhizobia in the early rhizosphere. It seems unlikely that

this early growth would comprise a mechanism for host-bacterium

specificity, but perhaps the competitive advantage of some strains over

others is a result of rapid early growth in the rhizosphere.

The basis for specificity in the Rhizobium-legume association

remains uncertain. In these studies, specific stimulation of Rhizobium

species in the rhizospheres of their homologous host plants was not

observed.

109

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